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PCR Testing, DNA Concentration Calculations,
Technical Log
Name: HaoQi (Esther) Li
Date: 7/20/06 Thursday
Work Location: Naval Medical Research Center, Silver Spring, MD
Title of Project: Developing and Optimizing a qPCR Assay for Borrelia lonestari
1. Objective – Why did you hope to accomplish today?
I plan to work out the concentration of the samples today and then do a dilution test standard.
Then I should start writing the paper.
2. Events – a. summarize the schedule of events. Procedures performed in detail, results,
analysis, with tables, charts, diagrams, and scanned sketches
b. summarize communications, emails (attached), conversations, pertinent material read
(annotated bibliography)+ reactions, browser used (search identifier, annotation)
I added 10%, i.e. 50μl of glycerol anhydrous to tubes #1, 3, and 5 in order to keep the bacteria
frozen alive. However, there was a problem with the viscosity of the glycerol anhydrous. I
started using 200μl of pipet, but it was so hard to extract it out so I used 1000μl and added more.
However, I doubt that I added exactly 50μl, but pretty close.
PCR Testing:
I did a PCR test for my five samples.
5 samples + 1 negative control + 1 extra = 7 samples to make
Date:
7/20/2006
Final conditions
Vol. Template
Vol. Reaction
Forward Primer
Reverse Primer
Reagent
Water
superMix 1.1X
11Forward Primer
970Reverse Primer
Specificity:B. lonestari
PCR Stage1:
1 ul
Stage2:
25 ul
0.3 uM
0.3 uM
Stage3:
Stock
22 ul/each
20 uM
20 uM
Total(Before Template Addition)
Each(Before Template Addition)
No.Samples 7
hold 94C: 2min.
3-Temp 36 cycles
94C: 30sec.
52C: 30sec
72C: 1min.
hold 72C: 7min.
Master Mix
8.74 ul
154 ul
2.63 ul
2.63 ul
168 ul
24 ul
I confused PCR with qPCR and brought the master mix to the dirty hood, but realized that the
little PCR tubes are in the clean room. Since I couldn’t bring the master mix back to the clean
room again, I had to start over and this time, aliquot the master mix in the clean room.
While loading the DNA samples (the samples after boiled for 10 minutes), the cap of #2 broke
off. So there might be possible contamination.
Then I used the red Tgradient PCR machine.
When I poured out the agarose gel solution, I realized that it was not enough so I made some
more gel solution following the directions of journal 7/12/06.
Then I loaded 8μl of samples (after mixing 8μl of sample with 2μl of dye) but 3.5μl of the
standard. My loading skills were very bad this time, and some dye came out of the well for
every sample, but the samples in the buffer did not matter because only the gel was imaged
under the UV.
Then I viewed the results under the UV light:
The image obtained clearly showed that all five samples contain DNA of about 1000 base pairs,
however, the quality of the run was just horrible: the ladder was slanted, the 100bp was very
much behind the first marker dye line, and the long trails ahead of the bands.
While removing the gel from the UV light, I broke it intentionally. However, I realized that
there were two layers! With much excitement, I took a shot at a piece of the top layer:
20-Jul-2006 17:00:53 Low=0 High=255 Gamma=1.0
Exposure=19.40secs
The left side of the image shows half of the wells because I broke the other half off. This did
not contain much of the standard lane with only 3.5μl of sample at the bottom, but it did clearly
show the unwanted trails of the samples.
Then I took a picture of the rest of the gel, with the bottom layer upwards and both layers
downwards (the above image fits like a puzzle with the image below):
Thus the double layer does hinder the results of the bands by making the two layers run at
different speeds (or maybe the touching layer run faster). So I took out the rest of the top layer
and obtained a beautiful image of the bottom layer only:
(Interesting note, the second to bottom lane looked like it contained the most loaded material
before I ran the gel, but it showed up the least under UV.)
So after all this, the PCR did verify that the samples from the bacteria cloning contained our
target B. lonestari gene sequence.
DNA Concentration Calculations:
From yesterday’s photometer results, Dr. Ju told me to do the dilution for sample #2, which has
an average concentration of 0.34μg/μl. I looked up the length of the bacteria vector in the
TOPO Cloning manual.
DNA Concentration (ug/ul)
0.34
DNA Length bp Vector: pCR-XL-TOPO
Insert: 970bp -10bp
total:
3519
960
4479
for dsDNA
1bp = 660 g/mol
Molecular Weight of the plasmid:
Number of 1 mol = 6.022 x 10^23 copies of DNA
1. Given that 1bp = 660 g/mol
(4479bp)(660g/mol /bp) = 29546140 g/mol, i.e. the molecular weight
2. Given that the DNA concentration is 0.34μl/μl:
(0.34μg/μl) (g/1000000μg) = 3.4 x 10-7 g/μl
3. (3.4 x 10-7g/μl) (mol / 2956140g) = 1.15 x 10-13 mo/μl
4. (1.15 x 10-13 mol/μl) (6.022 x 1023 copies / mol) ≈ 6.93 x 1010 copies/μl
In the same way, the number of copies for the average DNA concentration, 0.29μg/μl of sample
#4 is: (0.29μg/μl) (g/106μg) (mol/2956140g) (6.022x1023copies/mol) ≈ 5.91x1010copies/ul.
With reference of journal 7/07/06, I diluted DNA sample #2.
Start with 10μl of #2 6.93 x 1010copies/μl
Then add 59.3μl TE buffer since 69.3μl final volume - 10μl samples = 59.3μl TE buffer
Because, (10μl)(6.93x1010copies/ul) / (69.3μl final volume) = 1 x 1010 copies
Procedures for making about 500μl samples (modified from 7/07/06 log):
1. Mix 10μl of DNA sample #2 with 59.3μl of TE buffer  make 1010 copies of DNA
2. Mix 50μl of previous sample with 450μl TE buffer  dilute 1010 to 103 by 10’s
3. Mix 158.23μl of previous sample with 341.77μl TE buffer  dilute 103 to 100 by 100.5’s
Math: x μl + 2.16x μl = 500μl, thus x = 158.23μl
3. Reflections
a. actually accomplished: I did PCR testing to verify that the DNA cloned by bacteria is
indeed the target sequence. Then I calculated and performed DNA dilution.
b. concerns: Not today!
c. learning (workplace, science, project, yourself): I learned that making a double-layered
gel is not a good idea.
4. Planning
I hope to do the Optimization Testing for the primers. Then aliquot the bacteria from the 50ml
tubes into little cylindrical tubes. Then do more optimization tests.